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The Journal of Neuroscience, October 15, 1998, 18(20):8214-8227
Delayed Release of Neurotransmitter from Cerebellar Granule
Cells
Pradeep P.
Atluri and
Wade G.
Regehr
Department of Neurobiology, Harvard Medical School, Boston,
Massachusetts 02115
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ABSTRACT |
At fast chemical synapses the rapid release of neurotransmitter
that occurs within a few milliseconds of an action potential is
followed by a more sustained elevation of release probability, known as
delayed release. Here we characterize the role of calcium in delayed
release and test the hypothesis that facilitation and delayed release
share a common mechanism. Synapses between cerebellar granule cells and
their postsynaptic targets, stellate cells and Purkinje cells, were
studied in rat brain slices. Presynaptic calcium transients were
measured with calcium-sensitive fluorophores, and delayed release was
detected with whole-cell recordings. Calcium influx, presynaptic
calcium dynamics, and the number of stimulus pulses were altered to
assess their effect on delayed release and facilitation. Following
single stimuli, delayed release can be separated into two components:
one lasting for tens of milliseconds that is steeply calcium-dependent,
the other lasting for hundreds of milliseconds that is driven by low
levels of calcium with a nearly linear calcium dependence. The
amplitude, calcium dependence, and magnitude of delayed release do not
correspond to those of facilitation, indicating that these processes
are not simple reflections of a shared mechanism. The steep calcium
dependence of delayed release, combined with the large calcium
transients observed in these presynaptic terminals, suggests that for
physiological conditions delayed release provides a way for cells to
influence their postsynaptic targets long after their own action
potential activity has subsided.
Key words:
delayed release; asynchronous release; calcium; stellate
cell; granule cell; facilitation
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INTRODUCTION |
Action potential invasion of a fast
chemical synapse provokes a rapid, brief increase in the rate of
quantal release of neurotransmitter. At many types of synapses this
period of "phasic" release is followed by a much smaller but
longer-lived (tens of milliseconds to seconds) elevation of quantal
release rate called "delayed" release (Barrett and Stevens, 1972 ;
Rahamimoff and Yaari, 1973; Zengel and Magleby, 1981; Zucker and
Lara-Estrella, 1983; Cohen and Van der Kloot, 1986; Goda and Stevens,
1994; Van der Kloot and Molgo, 1994). Understanding the mechanism
responsible for delayed release (DR) may provide insight into the links
between presynaptic calcium, the time course of probability of release,
and various forms of use-dependent synaptic enhancement.
Paired-pulse facilitation (PPF), a form of synaptic enhancement that
lasts for hundreds of milliseconds after single action potentials
(Zucker, 1996), shares several features with DR, leading to the
hypothesis that they share a common underlying mechanism (Rahamimoff
and Yaari, 1973; Van der Kloot and Molgo, 1994). In support of this
hypothesis, facilitation and DR have been found to have similar time
courses at many synapses (Zengel and Magleby, 1981; Van Der Kloot and
Molgo, 1993; Goda and Stevens, 1994), and both phenomena are dependent
on residual presynaptic calcium (Van Der Kloot and Molgo, 1993; Kamiya
and Zucker, 1994; Atluri and Regehr, 1996; Cummings et al., 1996). The
mechanistic link between DR and facilitation is, however, controversial
(Zucker and Lara-Estrella, 1983; Van der Kloot and Molgo, 1994).
Another important issue is the calcium dependence of DR. High
concentrations of calcium chelators in presynaptic terminals can
eliminate DR (Van Der Kloot and Molgo, 1993; Cummings et al., 1996).
Furthermore, there is evidence that elevations in presynaptic calcium
levels increase miniature EPSC (mEPSC) frequency. For example, at the
crayfish neuromuscular junction, elevations in presynaptic calcium
produced by either sustained high-frequency stimulation or by calcium
ionophores increase mEPSC frequency (Delaney and Tank, 1994; Ravin et
al., 1997). The dependence of DR on intracellular calcium is not known
for synapses in the mammalian CNS, where it only recently has
become possible to measure presynaptic calcium levels.
Here we study the relationships between presynaptic calcium,
facilitation, and DR for synapses from cerebellar granule cells onto
stellate cells and Purkinje cells. In the cerebellar cortex, granule
cell axons rise from the granular to the molecular layer, bifurcate,
and run for several millimeters in either direction as so-called
"parallel fibers" (Palay and Chan-Palay, 1974). Parallel fibers
make en passant glutamatergic synapses with both Purkinje cells and small inhibitory interneurons, called stellate cells. Each of
these types of synapses has its own advantages: we already have studied
the role of calcium in facilitation at the synapse onto Purkinje cells
(Atluri and Regehr, 1996), whereas the stellate cell synapse is better
suited for detection of DR.
At both of these synapses we found that, after single stimuli, delayed
release can be separated into two components: a rapid component that
lasts for tens of milliseconds and a more sustained component that
lasts for hundreds of milliseconds. Although DR and facilitation are
both calcium-dependent, the amplitudes and time courses of these
components do not correspond to those of facilitation. A comparison of
presynaptic calcium levels and DR indicates that the rapid component of
DR is steeply calcium-dependent, whereas the sustained component is
driven by low levels of calcium with a nearly linear calcium
dependence. A simple model that incorporates a calcium-driven process
with slow kinetics is proposed to account for the calcium dependence of
DR.
Portions of this study have been published in abstract form (Atluri and
Regehr, 1997).
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MATERIALS AND METHODS |
Transverse cerebellar slices (300 µm thick) were cut from 13- to 17-d-old Sprague Dawley rats, as described previously (Llano et al.,
1991; Atluri and Regehr, 1996). Experiments were conducted at 23 ± 0.5°C, unless otherwise indicated. The control external solution
(2 ml/min flow rate) consisted of (in mM): 125 NaCl, 2.5 KCl, 2 CaCl2, 1 MgCl2, 26 NaHCO3, 1.25 NaH2PO4,
25 glucose, 0.02 bicuculline, and 0.05 D,L-2-amino-5-phosphonovalerate bubbled with 95%
O2/5% CO2. The low-calcium external
solution contained 1 mM CaCl2 and 2 mM MgCl2.
For EGTA-AM application experiments a 100 mM stock solution
of EGTA-AM in DMSO was aliquoted and frozen. Immediately before use,
the aliquots were diluted in external saline to final EGTA-AM concentrations consisting of 100, 20, 5, or 1 µM and DMSO
concentrations (by percentage) of 0.1, 0.02, 0.005, and 0.001. EGTA-AM
bath applications were 15 min long, with flow rates of 2 ml/min.
Detecting presynaptic calcium transients. Granule cell
axons, known as parallel fibers, and presynaptic terminals were labeled by local application of a solution containing the membrane-permeant forms of calcium indicators, as described previously (Regehr and Tank,
1991; Regehr and Atluri, 1995). The loading time was 8-10 min for
magnesium green-AM (Molecular Probes) (Zhao et al., 1996). Experiments
commenced 2 hr after dye loading. Parallel fiber tracts were stimulated
extracellularly with an electrode placed in the molecular layer near
the fill site. Fluorescence changes were measured in a
150-µm-diameter spot 400-700 µm away from the stimulus site. The
filter set used for magnesium green was a 450-490 nm excitation, an
FT510 dichroic, and an LP520 emission filter (Zeiss, Oberkochen,
Germany). Fluorescence was detected with a photodiode.
Measuring synaptic currents. Whole-cell recordings of
stellate cells (Barbour et al., 1994; Sabatini and Regehr, 1996) were obtained by using 2.0-3.0 M glass pipettes containing an internal solution of (in mM): 35 CsF, 100 CsCl, 10 EGTA, 10 HEPES,
0.1 D600, pH 7.3 with CsOH. Recordings of Purkinje cells used 1.0-2.0 M glass pipettes, with an internal solution of (in mM):
110 Cs2SO4, 10 EGTA, 4 CaCl2, 1.5 MgCl2, 5.5 MgSO4, 4 Na2-ATP, 0.1 D600, 5 QX-314,
and 10 HEPES, pH 7.3 with CsOH (Chen and Regehr, 1997). Stellate cells
were voltage-clamped at  40 mV and Purkinje cells at 70 mV. The
access resistance and leak current of stellate (from 0 to 20 pA) and
Purkinje (from 0 to 200 pA) cells were monitored continuously.
Parallel fibers were stimulated extracellularly with a glass
microelectrode placed in the molecular layer several hundred microns from the recording electrode. Stimulation rates were once per
7.5 sec or twice per 15 sec for delayed release and paired-pulse facilitation, respectively. These stimulation rates were chosen to
allow for the comparison of PPF with our previous study (Atluri and
Regehr, 1996). For all of our experimental conditions, by 2000 msec
after the conditioning pulse the PPF decayed to <10% (data not shown)
from a maximum of ~480% in 1 mM Ca and 190% in 2 mM Ca. The fast decay phase of the resulting EPSC was well
approximated by an exponential with a time constant of 1.5-3 msec for
stellate cells and 5-7 msec for Purkinje cells.
Measuring delayed release and facilitation. The first 10 msec of the EPSC, which is composed of many superimposed mEPSCs, was
not analyzed. Stellate cell DR was quantified by constructing peristimulus time histograms (bin width, 2 msec) of the occurrence of
mEPSCs. For summarizing data taken from several cells, it was necessary
to normalize for the number of stimulated fibers. After subtraction of
the prestimulus spontaneous mEPSC rate, histograms from each cell were
normalized by the number of events from 10 to 1000 msec and then
averaged together. For experiments in which cells were studied before
and after EGTA-AM or low calcium bath application, the postapplication
histogram was normalized by using values from the preapplication
histogram, preserving their relative amplitudes.
Facilitation was computed for each  t in two ways,
as 100 (A2 A1)/A1 and as
A2 A1, where A1 and
A2 were the integrated areas of the conditioning and test
EPSCs, respectively. These integrations were performed over a 20 msec
window. For t < 30 msec, the waveform of the
conditioning EPSC was subtracted from the superimposed test EPSC before
integration of the test EPSC waveform.
Experimental conditions were optimized for the detection of
facilitation and delayed release in the 10-1000 msec after
stimulation. Small stimulus intensities were used in the study of
facilitation to avoid series resistance artifacts. Larger stimulus
intensities generally were used to study delayed release to evoke
sufficient events in the 10-1000 msec range. There was no systematic
difference in the time course or relative amplitude of delayed release
on the basis of stimulus intensities. For large stimulus intensities it
was generally possible to detect all of the events 10 msec after
stimulation. However, it was not possible to detect all of the events
immediately after stimulation. For this reason, differences in the time
courses of EPSCs, such as those shown in Figures 3A and
4A, are best studied with much smaller stimulus intensities than those used in this study (C. Chen, P. Atluri, W. Regehr, unpublished observations).
Conditions of high release probability were avoided for two reasons.
First, we wished to avoid saturation of release. From previous studies
we know that, at this synapse, release is well approximated by a power
law, with n = 2.5 for external calcium concentration
(Cae) levels up to 2 mM (Mintz et al.,
1995), but that saturation is apparent when Cae is
>2.5 mM. Similarly, two-pulse DR experiments (see Fig. 8)
were conducted in 1 mM Cae to avoid saturation
of release. Second, for conditions of high release probability, such as
for three pulses in 2 mM Cae and for all experiments in higher Cae, the shape of the stellate
cell EPSC becomes prolonged; with sufficient numbers of pulses it can
depart from a single exponential decay, developing a plateau phase that can last for hundreds of milliseconds (P. Atluri, B. Sabatini, A. Carter, and W. Regehr, unpublished observations). This component reversed at ~0 mV and was completely blocked by either
6-cyano-7-nitroquinoxaline-2,3-dione or GYKI. It is likely a
consequence of spillover of neurotransmitter release. In the
experiments that were used in this work, such a component was not
present.
For practical reasons the experiments were not conducted in conditions
of extremely low release probability. As a consequence of the steep
calcium dependence of the first component of delayed release, we found
that, in 0.5 mM Cae, it was difficult to
obtain a sufficient number of events. We also performed a number of
experiments with BAPTA-AM, but these too were not suitable for
quantitative analysis. The strategy was to use this fast buffer to
reduce the size and slow the calcium transient and then to test the
effect on delayed release. Our model for DR predicted a large reduction in the number of events and a slowing of the decay of delayed release.
Qualitatively, this appeared to be the case. However, although BAPTA-AM
had the desired action on the calcium transient, there was
variability in the magnitude of this effect. Combined with the steep
calcium dependence of delayed release, this variability in the calcium
transient amplitude led to a much higher variability in the
magnitude of delayed release after loading with BAPTA-AM.
Data acquisition and analysis. Outputs of the photodiode and
Axopatch 200A were filtered at 500 and 2 kHz, respectively, with a
model 900C9L8L eight-pole Bessel filter (Frequency Devices, Haverhill,
MA) and digitized with a 16-bit D/A converter (Instrutech, Great Neck,
NY), Pulse Control software (Herrington and Bookman, 1995), and an
Apple Macintosh Centris 650 computer. Analysis was done on- and
off-line with Igor Pro software (WaveMetrics, Lake Oswego, OR).
Exponential fits to facilitation were performed with t
between 10 and 1000 msec. The amplitude of facilitation was calculated
from this fit for t = 10 msec. Double-exponential fits to calcium measurements and to delayed release histograms (see
Table 1) were performed between 10 and
1000 msec poststimulus, except as noted.
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RESULTS |
Delayed release at the granule to Purkinje cell and granule
to stellate cell synapses
Synaptic currents produced by single-pulse stimulation of parallel
fibers were recorded from both Purkinje cells and stellate cells.
Figure 1A1 shows the
average EPSC measured in a Purkinje cell, and 10 sequential trials from
this experiment are shown in Figure 1A2.
Corresponding recordings from a stellate cell are shown in Figure 1,
B1 and B2. As shown in Figure
1B3, the time course of release probability elevation
could be quantified by constructing a peristimulus time histogram of
mEPSC occurrence for the granule cell to stellate cell synapse.
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Figure 1.
Comparisons of phasic and delayed release at the
granule cell to Purkinje (A) and granule cell to
stellate (B) cell synapses evoked by 0.133 Hz
electrical stimulation of the parallel fibers. Shown are phasic EPSCs
(A1, B1) and 10 consecutive trials showing delayed
mEPSCs (A2, B2). These trials are blanked from 0 to 20 msec (A2) or from 0 to 10 msec (B2) for
clarity and are offset vertically by 200 and 100 pA, respectively. The
peristimulus time histogram (B3) of the stellate cell
mEPSCs is blanked from 0 to 10 msec. C, Semilogarithmic
plot of normalized representative Purkinje (thick line)
and stellate (thin line) cell EPSCs (same traces as
A1 and B1) and stellate cell mEPSC
histogram (dashed line). Traces in A1 and
B1 are averaged from 197 and 392 trials,
respectively. With an approximation of the decays of synaptic currents
with single exponentials, the time constants of decay for the average
mEPSC and evoked EPSC are 6.4 and 7.8 msec for the Purkinje cell in
A and 0.8 and 2.3 msec for the stellate cell in
B. The synaptic current measured in A is
somewhat slowed, because large stimulus strengths were used to
accentuate the delayed release of neurotransmitter. The spontaneous
mEPSC frequency was 9.9 Hz for the Purkinje cell in A
and 0.03 Hz for the stellate cell in B.
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A comparison of Figure 1, A and B, reveals some
obvious differences in the synaptic currents recorded in these two
cell types. The time courses of mEPSCs and evoked EPSCs are both much
faster in stellate cells than in Purkinje cells. This is consistent
with previous findings at these synapses and is thought to reflect the
prolonged presence of glutamate at the granule cell to Purkinje cell
synapses (Barbour et al., 1994). In addition, spontaneous mEPSC
frequencies are much higher in Purkinje cells than in stellate cells,
likely reflecting the relative number of synapses onto these two cell
types.
Despite these differences the timing of transmission from parallel
fibers onto these two postsynaptic targets is remarkably similar. At
both types of synapses the mEPSC frequency is elevated transiently
after the phasic EPSC, reflecting an increase in the probability of
release. Figure 1C, which compares the time courses of the
normalized average Purkinje and stellate cell EPSCs (Fig. 1A1,B1) and the normalized average
stellate cell mEPSC histogram (Fig. 1B3), also
suggests that delayed release is quantitatively similar in these two
types of cells. The rationale for using the EPSC as a measure of the
time course of delayed release [a similar approach has been used
previously (Goda and Stevens, 1994)] requires that (1) mEPSCs be
short-lived as compared with the time course of release probability,
(2) mEPSCs summate linearly, and (3) no other currents contribute to
the synaptic current. The good agreement between the stellate cell
histogram and EPSC shows that, for our conditions, stellate cell EPSCs
are accurate measures of the neurotransmitter release probability time
course. Also, the agreement between Purkinje and stellate cell EPSCs
strongly suggests that the granule cell presynaptic terminals that
synapse on Purkinje and stellate cells have very similar time courses
of release probability.
Because asynchrony of release can occur on rapid time scales (Katz and
Miledi, 1964; Diamond and Jahr, 1995; Isaacson and Walmsley, 1995),
identifying the transition between phasic and delayed release is
difficult. An operational distinction derives from the observation of
multiple components in the decay of release rate and from the
differential effects of pharmacological agents and gene knock-outs
(Miledi, 1966; Zengel and Magleby, 1980; Geppert et al., 1994). Here,
we study the quantal release of neurotransmitter in the 10 msec to 1 sec after stimulation of the presynaptic terminal. Although Figure
1C suggests that average EPSCs can provide a good measure of
the time course of delayed release in this time period, we feel that
the detection of individual events is more reliable. The ease with
which the very rapid mEPSCs can be discriminated, combined with the low
spontaneous mEPSC frequency, makes stellate cells ideal for the
detection of individual mEPSCs and for studies of delayed release.
Thus, for the remainder of the paper, we focus primarily on the
detection of individual mEPSCs and the study of delayed release and
facilitation in stellate cells.
Facilitation at the granule cell to stellate
cell synapse
The granule to stellate cell synapse demonstrates robust
paired-pulse facilitation (Fig. 2). The
inset of Figure 2 shows two EPSCs produced by stimulus pulses separated
by 40 msec. The second EPSC is much larger than the first, reflecting
an enhancement of the probability of glutamate release from granule
cell presynaptic terminals. This enhancement in evoked release
probability is transient, as shown in the main panel of Figure 2. In
this cell the peak amplitude of PPF was ~160%, and PPF decayed
monoexponentially with a time constant of 265 msec. For a series of
such experiments (n = 6), the amplitude of PPF was
181 ± 32% and the time constant of decay was 227 ± 27 msec, which is similar to that described previously at the granule cell
to Purkinje cell synapse [amplitude, 153 ± 11%;
tau, 203 ± 18 msec; n = 15 (Atluri and Regehr,
1996)].
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Figure 2.
Paired-pulse facilitation at the granule to
stellate cell synapse. The percentage of facilitation of integrated
EPSC areas, 100 (A2 A1)/A1, is plotted as a function of
interstimulus interval. Each point is the mean PPF ± SEM; n = 10. The smooth curve is
a monoexponential fit, omitting the first data point.
Inset, Synaptic currents evoked by extracellular
stimulation of the parallel fibers with pulses separated by 40 msec.
The shaded areas labeled A1 and
A2 illustrate the integration times that were used to
calculate the charge per pulse.
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A comparison of DR and PPF is a special case of the more general
problem of the relationship between mEPSC frequency and evoked EPSC
amplitude at some time. We define DR as the frequency of mEPSCs, which,
within an experiment, is proportional to the probability of release,
pr. PPF was computed in two ways:
A2 A1 and the more traditional measure,
100 (A2 A1)/A1. If we assume
that the number of release sites, n, is fixed, then the
values of A1 and A2 are proportional to
p(0) and p(t), the probabilities of evoked release in the instant before and at time t after
the conditioning pulse, respectively. A2 A1 is a measure proportional to the absolute
change in evoked release probability, p = p(t) p(0), whereas the
traditional measure of PPF is proportional to the relative
change in evoked release probability, p/p(0). As such, we feel that A2 A1 is better
suited for comparison to DR, although we also report the more
traditional measure of facilitation.
Manipulations of presynaptic calcium transients have
differential effects on facilitation and delayed release
A comparison of delayed release (see Fig.
1B) and facilitation (Fig. 2) at the granule cell to
stellate cell synapse suggested that these synaptic phenomena have
different time courses. This prompted us to explore further the calcium
dependence of these phenomena.
We tested the effects on DR of lowering Cae and altering
presynaptic calcium transients with EGTA, both of which have been shown
to affect facilitation (Atluri and Regehr, 1996). We took two
approaches to compare facilitation and delayed release. First, in some
experiments such as those shown in Figures
3 and 4,
facilitation and delayed release were both monitored. In such
experiments paired-pulse facilitation was monitored for a single
interpulse interval (t = 20 msec), using small
stimuli that would allow for an accurate determination of the magnitude
of facilitation. These trials were separated by 10 trials that used
single stimuli of a higher intensity to examine delayed release (see
Materials and Methods for more details). Second, a series of
experiments was conducted in which facilitation was measured for a
series of interpulse intervals for the same experimental conditions as
in Figures 3 and 4; these curves are summarized in Figure
5, where they are compared with the
delayed release histograms. Because it took many trials to obtain
facilitation curves and delayed release histograms, it was generally
not practical to obtain both sets of data for a cell in both
experimental conditions.
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Figure 3.
Differential effect of external calcium
concentration on facilitation and delayed release. Synaptic currents
were recorded in 2 mM Ca (control) and during bath
application of 1 mM Ca. Each double-pulse facilitation
trial (A) was followed by 10 single-pulse delayed
release trials (B). A, Areas of
conditioning (filled circles) and test
(open circles) EPSCs are plotted (left
panel) during bath application of 1 mM
external calcium. The right panel shows traces that are
averages of 14 trials for control (thick line) and low
calcium (thin line) on the same vertical scale
(top panel) or that are normalized (bottom
panel) to the peak of the first EPSC. There was a slight
decrease in the duration of the facilitated EPSC in 1 mM
Cae as compared with 2 mM Cae (see
Materials and Methods). B, Ten successive traces, each
offset vertically by 100 pA, in control (left
panel) and low (right panel)
calcium. C, Histograms of mEPSCs in control (left
panel) and in low calcium (right
panel). In B and C the
times from 0 to 10 msec have been blanked for clarity.
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Figure 4.
Differential effect of 20 µM EGTA-AM
on facilitation and delayed release. Double-pulse facilitation trials
(A) were interspersed among single-pulse delayed
release trials (B). A, Areas of
conditioning (filled circles) and test
(open circles) EPSCs are plotted (left
panel) during a 15 min bath application of 20 µM EGTA-AM. The right panel shows control
(thick) and postapplication (thin) traces
that are averages of 7 and 13 trials, respectively. The top
traces are on the same scale; the bottom traces
are normalized to the peak of the first EPSC. EGTA-AM decreased the
duration of the facilitated EPSC (see Materials and Methods).
B, Ten successive trials before (left
panel) and after (right panel)
treatment with EGTA-AM. C, Histograms of mEPSCs before
(left panel) and after (right
panel) EGTA-AM treatment. In B and
C the times from 0 to 10 msec have been blanked for
clarity.
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Figure 5.
Summary of the effect of low calcium and 20 µM EGTA-AM on facilitation and delayed release. The
average time course, from 0 to 300 msec, of A2 A1 facilitation (open circles) and
delayed release are plotted together for control (top),
low calcium (middle), and 20 µM
EGTA-AM-treated slices (bottom). Delayed release
histograms for low calcium and for 20 µM EGTA-AM first
were normalized by the associated pretreatment histograms from each
cell and then were averaged together.
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In Figure 3, Cae was lowered from 2 to 1 mM
while both facilitation and delayed release were monitored. Reducing
Cae decreased the peaks and areas of EPSCs evoked by
conditioning pulses and the test pulses. Facilitation, defined as
(A2 A1)/A1 or
A2 A1, increased by 111% or
decreased by 48%, respectively. There was an even greater
decrease in delayed release, as shown in the raw traces of Figure
3B and in the histograms of Figure 3C. In this
experiment the amplitude of delayed release at 20 msec poststimulus was
decreased by ~68%, but the time course of delayed release was not
altered significantly. [The experiment shown here is unusual in that
the spontaneous mEPSC rate was higher in 1 mM Ca. In a series of such experiments (n = 7) the spontaneous
mEPSC rate decreased from 3.4 ± 1.4 Hz in 2 mM Ca to
1.6 ± 0.6 Hz in 1 mM Ca.]
Another way of manipulating presynaptic calcium transients is to load
parallel fibers with EGTA (Atluri and Regehr, 1996; Feller et al.,
1996). The introduction of this slow but high-affinity calcium buffer
has small effects on the amplitude of the calcium transients but
greatly speeds the decay of residual calcium, [Ca]i. The
introduction of EGTA does not alter significantly the resting calcium
level (which is set by the balance of influx vs efflux; see Tank et
al., 1995). By the use of a membrane-permeant form of EGTA (EGTA-AM),
it is possible to introduce high concentrations of EGTA into
presynaptic terminals (free EGTA levels in the presynaptic terminals
can be much higher than the extracellular concentration of EGTA-AM).
Application of the membrane-permeant calcium chelator EGTA-AM has been
shown to alter the time courses of presynaptic calcium transients in
parallel fibers in a dose-dependent manner (Atluri and Regehr, 1996).
High concentrations of EGTA-AM also have been shown to eliminate
delayed release at other synapses (Cummings et al., 1996). In Figure 4,
facilitation and delayed release were measured before and after the
application of 20 µM EGTA-AM. EGTA-AM slightly decreased
the amplitudes of both the conditioning and the test pulses, and
reduced (A2 A1)/A1
facilitation by 39%, A2 A1 facilitation
by 71%, and the amplitude of delayed release (at 20 msec) by 96%
(Fig. 4B,C).
From Figures 3 and 4 it is apparent that manipulations of calcium entry
and calcium dynamics had differential effects on facilitation and DR.
The traditional measure of facilitation, (A2 A1)/A1, tended to overemphasize these
differences, but they remained even when A2 A1 facilitation was used. Figure 5 summarizes the results of
a series of experiments measuring facilitation and delayed release in
control conditions (top panel), in low external calcium (middle panel),
and after treatment with 20 µM EGTA-AM (bottom panel).
Each panel shows a normalized plot of A2 A1 facilitation and delayed release. In all cases the time
courses of decay of facilitation (control, 223 msec; 1 mM
Ca, 141 msec; EGTA, 55 msec) are slower than those of delayed release
(see Table 1), suggesting that the two processes are not identical.
The calcium dependence of delayed release from granule cell
presynaptic terminals
The manipulations of presynaptic calcium levels that are presented
in Figures 3 and 4 established the importance of presynaptic calcium to
delayed release at this synapse. To study directly how delayed release
depends on presynaptic residual free calcium concentration in parallel
fibers ([Ca]i), we measured [Ca]i
(Regehr and Atluri, 1995; Atluri and Regehr, 1996). Several
manipulations were used to alter the [Ca]i transient
evoked by single stimuli: (1) external calcium levels
(Cae) were reduced, (2) presynaptic Ca buffering was
altered, and (3) the number of action potentials was increased. During
these manipulations we monitored the amplitudes and time courses of
[Ca]i and delayed release.
We used the low-affinity calcium-sensitive indicator magnesium green
(Zhao et al., 1996) and previously described methods to measure
[Ca]i (Regehr and Atluri, 1995). The normalized change in
the fluorescence of magnesium green
(F/F) is directly proportional to
changes in [Ca]i and can be used to measure the time
course of presynaptic calcium transients and to detect changes in
calcium entry. Although these methods do not provide a reliable measure of the brief, highly localized calcium signals that drive phasic release (Fogelson and Zucker, 1985; Simon and Llinás, 1985;
Roberts, 1993), they are well suited to measuring [Ca]i
on a time scale relevant to delayed release. After an action potential,
calcium-binding proteins rapidly chelate Ca as it spatially
equilibrates by diffusion throughout the terminal. Provided that the
kinetics of Ca-binding proteins are rapid and that the presynaptic
boutons are small, spatial and chemical equilibration can occur within
several milliseconds (Connor and Nikolakopoulou, 1982; Sala and
Hernandez-Cruz, 1990; Roberts, 1994; Sabatini and Regehr, 1998). After
equilibration, average nerve terminal free Ca levels are tens to
hundreds of nanomolars above resting levels and may be detected by
fluorescent dyes. Then Ca is extruded from the terminal and returns to
resting levels over a period of hundreds of milliseconds.
Lowering external calcium decreases the size of the
F/F signal, as shown in a representative
experiment in Figure
6A. The left panel
shows the reversible effect of lowering Cae from 2 to 1 mM on peak F/F transients. The
right panel shows F/F transients before and
during the 1 mM Cae bath application. In this
experiment the peak decreased by 40%, although the time course
remained unchanged.
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Figure 6.
The effect of external Ca concentration on
presynaptic Ca transients and delayed release. Fluorescence
(F/F) transients were monitored
in granule cell presynaptic terminals by the low-affinity Ca dye
magnesium green. A, Circles (left
panel) represent peaks of fluorescence transients during
a change in the bath solution from 2 mM Cae
(right panel, top trace) to 1 mM Cae (right panel, bottom
trace). B, Normalized average calcium transients
(thick traces) and delayed release histograms
(thin traces) are shown in 2 mM
Cae (top panel) and 1 mM
Cae (bottom panel),
respectively.
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Figure 6B compares the average time courses of
[Ca]i and delayed release for control conditions (top
panel) and in low calcium (bottom panel). In both conditions
[Ca]i and delayed release could be approximated by
double-exponential decays, and in both control and low calcium the
early and late components of release probability both decayed much more
rapidly than did [Ca]i (see Table 1). Lowering external
Ca decreased the amplitudes of [Ca]i and delayed release
by 38 and 75%, respectively (measured at 20 msec), without significantly altering their time courses.
We took advantage of the effects of EGTA on the calcium transient to
examine the relationship between the time course of [Ca]i and that of neurotransmitter release probability. In Figure
7A, we show a representative
experiment in which 5 µM EGTA-AM was applied to a slice
for 15 min. EGTA decreased the peak of the F/F
transient by 11% and decreased its half-decay time from 40 to 17 msec.
Treatment with different concentrations of EGTA-AM led to a
dose-dependent speeding of decay times [Fig. 7B, top panel; 0, 1, 20, and 100 µM traces replotted
from Atluri and Regehr (1996)]. Release probability histograms after
EGTA-AM treatment revealed a diminution of both the amplitude and
duration of delayed release. A direct comparison of calcium and delayed
release for various EGTA-AM concentrations showed that in all cases the
release probability declined more rapidly than does calcium (Fig.
7C, Table 1). The introduction of very high concentrations
of EGTA (in the 100 µM EGTA-AM experiments) eliminated
delayed release.
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Figure 7.
The effect of EGTA-AM concentration on presynaptic
Ca transients and delayed release. Fluorescence
(F/F) transients were monitored
as in Figure 6. A, Filled and open
circles (left panel) represent peaks and
half-decay times, respectively, of fluorescence transients before
(right panel, top trace) and after
(right panel, bottom trace) a 15 min
application of 5 µM EGTA-AM. B,
Application of 0, 1, 5, 20, or 100 µM EGTA-AM causes a
dose-dependent diminution and acceleration of normalized average
presynaptic Ca transients (top panel) and delayed
release histograms (bottom panel).
C, Comparison of normalized average calcium transients
(top traces) and delayed release histograms from 0, 1 5, 20, and 100 µM EGTA-AM experiments.
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We also examined the effect of the number of stimuli on the delayed
release of neurotransmitter. We measured delayed release or magnesium
green fluorescence after a single stimulus and after two stimuli
separated by 10 msec. These experiments were performed in 1 mM Cae to lower the initial probability of
release sufficiently to reduce the depression of transmitter release
after a double stimulus, which could complicate the interpretation of
such experiments if they were performed in 2 mM
Cae. Figure
8A shows a
representative experiment. There is clear facilitation of phasic
transmitter release in response to the second of two stimulus pulses,
as shown in Figure 8A1. In addition, after two pulses
there is a large increase in DR (compare Fig. 8A2
with A3). In this experiment there was a 185% increase in
the total number of events (from 20 to 1000 msec after the final pulse)
for the double-pulse trials as compared with the single-pulse trials.
The time course of delayed release was significantly slower after two
pulses than after a single pulse, whereas the time course of calcium
was only slightly slower after two pulses than after a single pulse
(see Table 1). As shown in Figure 8B, DR decays more
rapidly than does calcium after either single or double pulses.
However, increasing the pulse number has a much larger effect on the 20 msec amplitude of delayed release (which increased by 369%) than on
the 20 msec amplitude of calcium (which increased by 88%).
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Figure 8.
The effect of the number of stimulus pulses on
presynaptic Ca transients and delayed release in 1 mM
external Ca. Shown are phasic EPSCs (A1), delayed mEPSCs
(A2) in 10 consecutive trials, and mEPSC histograms
(A3) for single-pulse (left panels) or
double-pulse (right panels) stimuli. Times from 0 to 10 msec (left panels) or from 0 to 20 msec (right
panels) are blanked for clarity (A2,
A3). B, Normalized average calcium
transients (top traces) and delayed release histograms
after single (top panel) or double (bottom
panel) pulses.
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Modeling delayed release
We investigated several approaches to modeling the calcium
dependence of delayed release. Initially, DR was assumed to be a
function of the calcium, [Ca]i, that was present
in the presynaptic terminal,
![DR=A[<UP>Ca</UP>]<SUB><UP>i</UP></SUB>+B[<UP>Ca</UP>]<SUB><UP>i</UP></SUB><SUP>n</SUP>,](/content/vol18/issue20/fulltext/8214/img001.gif)
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(1)
|
where A, B, and n are
constants. This equation has been shown previously to describe the
relationship between calcium and the frequency of quantal release of
neurotransmitter at the crayfish neuromuscular junction (Delaney and
Tank, 1994; Ravin et al., 1997). We found that, for each of the
experimental conditions in this study, Equation 1 could provide a good
approximation to the observed delayed release (data not shown).
However, if delayed release is a function of intracellular calcium
levels, a single set of parameters should describe the relationship
between calcium and delayed release for all of the conditions (Figs.
6-8). This is not the case. As illustrated in Figure
9, curves relating DR to internal calcium
levels measured in 1 mM Cae (open circles) and
2 mM Cae (filled circles) do not superimpose.
For A = 0.08, B = 1.0, and
n = 3.5, this model overpredicts delayed release for 2 mM Cae and underpredicts delayed release for 1 mM Cae (Fig. 9A). Furthermore, no
single set of parameters provides a good description of delayed release
for the different external calcium conditions. The reason for this is
apparent in Figure 9B, in which the delayed release is
plotted as a function of the calcium concentration for 1 and 2 mM Cae. Although for low [Ca]i
these curves superimpose, for high [Ca]i they deviate
markedly. Models that assume delayed release to be a function of
calcium that is present in the terminals at that instant predict that
all curves relating delayed release and calcium should superimpose;
such models cannot account for the relationship between delayed release
and calcium in Figure 9B. In Figure
10 we also plot DR as a function of
[Ca]i for experiments varying external Ca (top panels),
pulse number (middle panels), or EGTA-AM treatment (bottom panels).
These DR versus Ca curves do not superimpose, and the differences are
pronounced in experiments with one and two pulses (middle panels).
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Figure 9.
Steady-state models relating calcium to delayed
release are inadequate. A, Normalized delayed release
versus time in the presence of 1 mM Cae
(open circles) and 2 mM Cae
(filled circles). Solid lines were
computed by using Equation 1, with A = 0.08, B = 1.0, and n = 3.5, and
calcium transients were measured experimentally (normalized to the
calcium value at 10 msec in control conditions). B,
Delayed release versus normalized [Ca]i for 1 mM Cae (open circles) and 2 mM Cae (filled circles).
The solid line is the computed delayed release versus
the normalized calcium transient from A. The curves that
were calculated for 1 and 2 mM Cae
overlap.
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Figure 10.
Summary plots of delayed release versus
presynaptic calcium levels after parallel fiber stimulation. Shown is
normalized delayed release as a function of time (left
panels) and as a function of calcium concentration
(right panels) in 1 and 2 mM Cae
(top), for one and two pulses in 1 mM
Cae (middle), and after loading with 1, 5, and 20 µM EGTA-AM (bottom). Graphs to the
right correspond to graphs on the left,
and the same symbols are used. In the bottom right graph
the 20 µM EGTA-AM graph was not included for clarity.
Solid curves are computed according to Equation 2, as
described in Results.
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On the basis of what is known about the timing of transmitter release
and the role of calcium in use-dependent processes (Stanley, 1986;
Yamada and Zucker, 1992; Regehr et al., 1994; Atluri and Regehr, 1996;
Bertram et al., 1996), it is not surprising that the assumption of
calcium instantaneously affecting the rate of delayed release appears
to be an oversimplification. The most straightforward way to account
for the relationship between calcium and DR for different experimental
conditions (Figs. 9B, 10) is to include the time dependence
of a calcium-driven process:
![DR=A[<UP>Ca</UP>]<SUB><UP>i</UP></SUB>+B·<UP>exp</UP>(<BINOM><NU><UP>−t</UP></NU><DE><UP>&tgr;</UP></DE></BINOM>)[<UP>Ca</UP>]<SUB><UP>i</UP></SUB><SUP>n</SUP>,](/content/vol18/issue20/fulltext/8214/img002.gif)
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(2)
|
with A, B,  , and n
constants. As in Equation 1, delayed release is made up of two
components. However, the second component is now both time-dependent
and calcium-dependent. The term represents the remnant of a process
with slow kinetics that is driven by high local calcium levels
immediately after action potential invasion of the presynaptic
terminals. The calcium-dependent term,
[Ca]in, corresponds to
multiple calcium ions binding to sites that respond rapidly to changes
in calcium levels and that must all be occupied for release to occur.
Although this model posits two additive components of delayed release,
it is not known whether they rely on distinct molecular mechanisms or
whether they share some of the same molecular machinery.
As shown in Figure 10, fits based on this simple scheme provide a good
description of the relationship between calcium and the rate of delayed
release for a variety of experimental conditions. The symbols in Figure
10 show the normalized delayed release as a function of time (left) or
as a function of normalized calcium concentration (right) after
stimulation by one or two pulses in a variety of experimental
conditions. The lines are values of DR calculated with Equation 2,
using the calcium transient measured for the corresponding experimental
conditions (normalized to the value measured in control conditions at
10 msec in 2 mM Cae). The same
parameters were used for all simulations (A = 0.1, B = 1.25, = 28 msec; n = 2.5). Thus
it is possible to relate [Ca]i and delayed release with a
simple model that incorporates the time dependence of release.
Delayed release at higher temperatures
Experiments described thus far have been directed toward
characterizing the properties of delayed release at this synapse and in
particular the role of calcium in delayed release. These studies were
performed at 23°C, owing to the ease of experimentation at room
temperature. We also investigated the properties of delayed release at
closer to physiological temperatures. As shown in Figure 11, delayed release at 33°C was
qualitatively similar to that observed at 23°C, although it was
somewhat shorter-lived. As at room temperature there was a large
increase in the magnitude of delayed release for two pulses compared
with that produced by a single pulse. Although these experiments were
still performed at lower than physiological temperatures, on the basis
of the temperature dependence of the magnitude and time course of
delayed release it is reasonable to assume that delayed release of
neurotransmitter from cerebellar granule cell terminals is also
prominent at physiological temperatures.
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Figure 11.
Delayed release of neurotransmitter that follows
one (left panel) and two (right
panel) pulses at 33°C. Shown are phasic EPSCs
(top), delayed mEPSCs (middle) in 10 consecutive trials, and mEPSC histograms (bottom). For
delayed release trials and for mEPSC histograms the times from 0 to 10 msec (right) or from 0 to 20 msec (left)
have been blanked for clarity.
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DISCUSSION |
Comparison of facilitation and delayed release
We tested the hypothesis that facilitation and delayed release
share a common underlying mechanism. Both components of DR decayed more
rapidly than did facilitation (see Figs. 1, 2, 5). Previous studies of
other synapses have reported similar mismatches between the decay of
quantal event frequency and the decay of peak release amplitude; in all
cases the quantal event frequency decayed more rapidly than did the
enhancement of evoked release (Zucker and Lara-Estrella, 1983; Van der
Kloot and Molgo, 1994). We also found that DR and PPF differed in their
responses to manipulations of presynaptic calcium influx (see Figs. 3,
5) and of calcium decay kinetics (see Figs. 4, 5). DR is more sensitive
to external calcium concentration than is facilitation. Accelerating
the decay of [Ca]i with EGTA-AM (see Appendix of Atluri
and Regehr, 1996) sped the decays of both DR and PPF (see Figs. 4, 5),
but the initial amplitude of PPF was decreased only slightly, whereas
that of DR was markedly reduced. Together, these findings suggest that facilitation and DR are not simple reflections of a common underlying mechanism at granule cell presynaptic terminals.
Calcium dependence of delayed release
A number of manipulations of presynaptic calcium transients
provided insight into the [Ca]i dependence of DR.
Treatment with high concentrations of EGTA-AM virtually eliminated DR,
confirming the calcium dependence of this process (Cummings et al.,
1996). Lower concentrations of EGTA-AM sped the decay of both
[Ca]i and DR in a dose-dependent manner (see Fig. 7).
Even modest concentrations of EGTA-AM, which did not affect the
amplitude of calcium significantly at early times, had a profound
impact on DR. Furthermore, DR was highly sensitive to manipulations of
external calcium concentration, suggesting that the bulk of DR was a
consequence of calcium ions acting cooperatively (see Fig. 6).
When DR was plotted as a function of [Ca]i, it
appeared to consist of two components, one linearly related to
[Ca]i and the other more steeply dependent on
[Ca]i (see Fig. 10). Steady-state models in which the
frequency of DR was a function of the instantaneous free Ca
concentration could not account for the relationship between [Ca]i and DR for all of the conditions of our
experiments. This deviation indicates that DR is not solely a function
of instantaneous Ca levels.
Incorporating kinetics into the model of DR, as in Equation 2,
recognizes that calcium-dependent processes do not necessarily reflect
instantaneous calcium levels. This has been shown previously for
calcium-driven phasic release (Katz and Miledi, 1964) and for
calcium-driven synaptic plasticity (Regehr et al., 1994; Atluri and
Regehr, 1996; Zucker, 1996), so it is reasonable to think that
time-dependent processes also would influence DR. The simple model we
used to relate [Ca]i and DR consists of two components that can be thought of as corresponding to calcium-binding sites with
different kinetics and affinities, much like those introduced in
previous models of synaptic transmission (Yamada and Zucker, 1992;
Bertram et al., 1996). According to such models, release is
proportional to the occupancy of multiple binding sites with different
properties. Undoubtedly, many other schemes also could relate
[Ca]i and DR in our experiments, but this extremely
simple model is useful in that it illustrates the ability of a model that incorporates kinetics to relate [Ca]i and DR. In
addition, it is clear that the high-power law relationship relating the DR and calcium curves for nonsteady-state conditions must be
interpreted with caution.
Our findings help to explain the properties of DR at various synapses,
which to a large extent may reflect the calcium dynamics of different
types of presynaptic terminals. The magnitude of DR that we observe for
one or two pulses applied at high frequency is remarkable. This may
reflect the large residual calcium transients that are present in these
tiny presynaptic terminals. At many other synapses DR is not so
prominent after such a small number of pulses. For example, in
crayfish, in which little DR is observed after single stimuli, it is
thought that single action potentials increase [Ca]i by
tens of nanomolars (Tank et al., 1995). In contrast, single action
potentials increase [Ca]i by several hundred nanomolars in granule cell presynaptic terminals (Regehr and Atluri, 1995).
Implications for molecular mechanisms of
delayed release
It has been suggested previously that two kinetic components of
delayed release at hippocampal synapses might reflect distinct mechanisms involving different molecules (Geppert et al., 1994; Goda
and Stevens, 1994). Our observation that two components of DR differ in
their calcium dependence supports this hypothesis and constrains the
properties of the molecular mechanisms. On the basis of our findings we
predict that the binding of a single calcium ion to a binding site
triggers one component of DR. We also predict that there is a component
of release that is driven by the cooperative binding of multiple
calcium ions and that multiple types of binding sites are likely to be
involved. This component of DR has many properties in common with
phasic release, and it likely involves many of the same molecules,
including synaptotagmin I (Geppert et al., 1994). Our findings also
suggest that different molecules may be involved in DR and
facilitation. It is difficult, however, to eliminate the possibility
that the same molecules are involved, yet they have differential
effects on spontaneous and evoked release. The calcium dependence of
the processes observed here should be helpful in future studies to
determine which of the many candidate molecules (Broadie et al., 1995;
Zucker, 1996; Wu and Bellen, 1997; Deitcher et al., 1998) are involved
in facilitation and in the two components of DR.
Physiological significance of delayed release
of neurotransmitter
Most investigations of delayed release have centered on the
insight that this phenomenon can provide about synaptic
transmission there has been very little consideration given to the
physiological importance of this phenomenon. This is not surprising in
considering the DR after a single impulse, as in Figure 1, which shows
that the vast majority of synaptic current occurs immediately after a
single stimulation, and the sporadic DR of neurotransmitter appears to
be insignificant by comparison. Normally, however, neurons do not fire
single impulses widely separated in time. More typically, firing rates
are tens of Hertz, and neurons often fire in high-frequency bursts
(Livingstone et al., 1996; Lisman, 1997). The response to such a burst
is revealing. In addition to the large, rapid component of release, two
stimulus pulses at 100 Hz produce a barrage of miniature synaptic
currents. Over four times as many mEPSCs are observed after two stimuli
as are seen after a single stimulus. This sensitivity to pulse number continues for three stimuli, where the integrated synaptic currents for
DR actually can be larger than the phasic component of release (P. Atluri and W. Regehr, unpublished observations). It seems likely that
DR will influence firing of the postsynaptic cell for hundreds of
milliseconds after a stimulus burst of the presynaptic inputs. This
suggests the exciting possibility that DR may provide a synapse with a
novel type of "short-term memory" that may have important
physiological consequences.
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FOOTNOTES |
Received June 8, 1998; revised July 28, 1998; accepted Aug. 5, 1998.
This work was supported by National Institutes of Health Grant
R01-NS32405-01. We thank A. Carter, C. Chen, J. Dittman, M. Friedman,
A. Kreitzer, and B. Sabatini for comments on this manuscript.
Correspondence should be addressed to Dr. Wade Regehr, Department of
Neurobiology, Harvard Medical School, 220 Longwood Avenue, Boston, MA
02115.
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Top
Abstract
Introduction
